2Present address: Kolling Institute of Medical Research,
Kolling Building, Royal North Shore Hospital, St. Leonards, New South
Wales, 2065, Australia

* To whom correspondence should be addressed.

Received November 29, 2010; Revision received February 2, 2011
Identifying differences in DNA methylation is critical to understanding
how epigenetics influences gene expression during processes such as
development. Here, we propose a method that employs a single,
methylation-sensitive restriction endonuclease of choice, to produce
discrete pools of methylated and unmethylated DNA from the same sample.
A pool of restriction fragments representing unmethylated regions of
the genome is first obtained by digestion with a methylation-sensitive
endonuclease. The restriction-digested DNA is then concatamerized in
the presence of stuffer-adaptor DNA, which prevents interference from
originally unmethylated DNA by blocking the ends of the restriction
fragments. The concatamerized DNA is amplified by phi29 polymerase to
remove methylation marks, and again digested with the same endonuclease
to produce a pool of DNA fragments representing methylated portions of
the genome. The two pools of DNA fragments thus obtained can be
analyzed by end-sequencing or hybridization to a genomic array. In this
report we detail a proof of concept experiment that demonstrates the
feasibility of our method.
KEY WORDS: whole genome methylation, phi29 polymerase, whole
genome amplification, next-generation sequencing

DOI: 10.1134/S0006297911090021

Profiling differences in DNA methylation across a genome is relevant to
understanding how epigenetic effects influence cell- and
tissue-specific gene expression and processes such as cancer [1, 2]. Use of isoschizomers that
possess differences in methylation-sensitivity, such as HpaII
and MspI, to probe differences in methylation is well
documented. With this approach, restriction fragments released after
digestion by HpaII are representative of unmethylated regions of
the genome, whereas an absence of HpaII digestion is diagnostic
for DNA methylation. Combined with next-generation sequencing, this
strategy was recently used to profile whole genome methylation using a
method dubbed as Methyl-Seq [3]. However,
methylation features are rarely unambiguous, and as such, a certain
proportion of DNA molecules at a particular locus will produce
fragments after digestion with methylation-sensitive endonucleases,
even for generally methylated regions, as has been shown with
particular epigenetic features [4]. While elegant
solutions to the digital measurement of methylation frequency exist,
such as MethylC-Seq [5] or BS-Seq [6], which employ bisulfite conversion [7] coupled to next-generation sequencing (e.g.
Illumina Genome Analyzer) — their cost, especially for large
genomes, may be prohibitive for some researchers [8].

Our solution to this problem is to use a single, methylation-sensitive
restriction endonuclease of choice to derive two discrete pools of
genomic fragments from the same sample — one containing
unmethylated loci and the other containing methylated loci. The
abundance of each fragment in these pools, and genomic location, could
be determined through end sequencing. It would also be possible to use
the fragments so obtained for hybridization to a whole genome
microarray or tiling array. Our method is advantageous, as it can
reduce the amount of sequencing required by methods such as
MethylC-Seq, while also being applicable to species without sequenced
genomes. The method we detail here should thus provide the ability to
scan any genome at different levels of granularity (depending on the
digestion frequency of the restriction endonuclease used) to identify
and quantify the prevalence of novel methylated loci.

Our method exploits phi29 polymerase, an enzyme demonstrating
unparalleled processivity and an ability to amplify long fragments of
DNA devoid of methylation, which are thus suitable for digestion by
methylation-sensitive restriction endonucleases [9]. Coupled to the ability of phi29 to amplify a
fragmented genome following concatamerization with short adaptors,
coined as stuffer DNA by Shoaib and coauthors [10], these features form the basis for a method that
permits the quantitative analysis of methylation features in whole
genomes. In our approach, DNA representing unmethylated regions of the
genome is first digested with a methylation-sensitive restriction
endonuclease. Next, methylation marks from methylated regions are
removed by amplification with phi29 polymerase. Amplified DNA is then
digested with the same endonuclease releasing originally methylated
fragments. Interference from originally unmethylated DNA is avoided by
blocking the ends of the restriction fragments with
specifically-designed, stuffer-adaptor DNA. Thus, two pools of DNA are
obtained that could be analyzed by end-sequencing or hybridization to a
genomic array. In this report, we detail a proof of concept experiment
that demonstrates the feasibility of our conceptual approach using an
artificially constructed pseudogenome.

MATERIALS AND METHODS

Synthesis of a pseudogenome. We synthesized a pseudogenome by
ligating a mixture of digested λ-DNA and linearized plasmid to
simulate unmethylated and methylated fractions of a genome,
respectively. To create the plasmid, we cloned a 7.3 kb Medicago
truncatula (cultivar Jemalong) genome fragment corresponding to
bases 36,490,391-36,497,718 of chromosome 4 (see Supplement (Fig. S1) on site of Biochemistry (Moscow) journal
http://protein.bio.msu.ru/biokhimiya). The genomic
fragment was amplified using the primers
5′-AACACCATTAGAAGCTTCTAGAATCGGAA-3′ and
5′-CATCGATGCAGTCGTATAAGTTAGTACTAG-3′ using the Expand Long
Template PCR System (Roche, Germany). The genome fragment contained two
ClaI sites, while the second primer introduced a third
ClaI site. The resulting PCR amplicon was cloned by TA cloning
into the pGEM-T vector (Promega, USA). Unmethylated λ-DNA (20
µg; Promega; D1521) and the plasmid (40 µg) were digested
by 20 units of ClaI (New England Biolabs, USA) in
100 µl reactions for 2 and 16 h at 37°C, respectively.
The ClaI was inactivated at 65°C for 20 min. The
efficiency of digestion was checked by gel electrophoresis.
Electrophoresis was performed using a 0.6% (w/v) agarose gel prepared
in Tris-acetate-EDTA buffer (pH 8.2) and run at 100 V, 22°C.
The λ-DNA (1 µg in a 5 µl volume) and
linearized plasmid (2 µg in a 5 µl volume) were
combined with 1.2 µl of 10× T4 DNA ligase buffer and
ligated by adding 1 µl of concentrated T4 DNA ligase (2000
cohesive end units; New England Biolabs, USA) for 24 h at
14°C.

Phi29 amplification and digestion. The stuffer-pseudogenome
ligation mixture (1 µl) was diluted to 5 µl with
water and used for phi29 amplification. The DNA was denatured at
95°C for 3 min and immediately chilled on ice. The
amplification reaction was performed in a total volume of
20 µl using the REPLI-g DNA polymerase and buffer from the
QuantiTect Whole Transcriptome Kit (Qiagen, Germany) at 30°C for
2 h and the reaction terminated at 65°C for 10 min. It
would be possible to amplify using phi29 from other suppliers. In such
cases, the reaction should be supplied with deoxyribonucleotide
triphosphates (1-4 mM) and random hexamers (5-50 µM). Addition of
pyrophosphatase and the use of modified (exonuclease-resistant) random
hexamers could potentially increase yields significantly [11]. In our hands, the reaction yielded approximately
20 µg of the amplified DNA. After amplification, the
reaction mixture was diluted to 100 µl with ClaI
buffer and digested with ClaI as described above.

RESULTS AND DISCUSSION

We used a mixture of λ-DNA and a plasmid containing a 7.3 kb
M. truncatula genome fragment to simulate unmethylated and
methylated fractions of a genome, respectively. We chose to use a
pseudogenome to more clearly illustrate how our procedure operates, but
we believe our method has utility to genuine genomes. Although genuine
genomes are longer and will exhibit more complex methylation profiles,
we believe that our simple pseudogenome accurately reflects how the
basic tenets of our procedure would operate on a genuine genome.

To produce the pseudogenome, we digested both the λ-DNA and
plasmid with the methylation-sensitive endonuclease ClaI and
then ligated the products of these digestion reactions (Fig. 1; see color insert). In this pseudogenome, the
ClaI-digested λ-DNA fragments represent completely
unmethylated genomic DNA (Fig. 1, blue regions;
Fig. 2, lane 2) whereas the digested plasmid
represents a region of methylated genomic DNA (~6 kb; see Supplement,
Fig. S1) bounded by the remnants of an
unmethylated ClaI site (Fig. 1, black
region; Fig. 2, lane 3). Note that although
the plasmid had three ClaI sites, two sites within the genome
fragment were selected such that they would be protected by bacterial
DNA adenine methylation (ClaI sites in a GATC context) and hence
ClaI digestion resulted in plasmid linearization. We favored the
use of in vivo adenine methylation over in vitro CpG
methylation as it is more efficient.

Subsequent digestion of the pseudogenome by ClaI, which results
in restriction fragments possessing CG overhangs, produced two types of
restriction fragment — predominantly short fragments arising from
unmethylated loci and longer fragments from methylated, and therefore
protected, loci. Although a range of fragment sizes would result from
digestion of a complex genome, the use of a frequent cutting
endonuclease would produce a majority of small (<2 kb) fragments. At
this point in the procedure, the short fragments representative of
unmethylated loci would be saved for further analysis by sequencing or
hybridization to a genomic array (Fig. 1). We
envisage that the loci contained in the short fragment pool would
enable a scan of most unmethylated loci in the genome. It would,
however, be possible to sequence the entire pool of restriction
fragments is so desired.

To release fragments of methylated DNA, the digested pseudogenome was
first concatamerized with an excess of stuffer-adaptor DNA (with
ClaI-compatible ends) before being amplified with phi29
polymerase (Fig. 1). The phosphorylated CG
overhangs of the ClaI-digested pseudogenome served to facilitate
ligation of stuffer-adaptor DNA, in turn enabling amplification of the
concatamerized pseudogenome by phi29. Ligation of stuffer-adaptor DNA
into the ClaI-digested pseudogenome acts to block the
re-digestion of unmethylated ClaI sites, while phi29
amplification removes methylation marks from originally methylated
ClaI sites. Hence, after digestion of the phi29-amplified DNA by
ClaI, fragments originally protected by methylation are released
and can be saved for further analysis (Fig. 1).

We prepared the stuffer-adaptor DNA by digesting a pCR8/GW/TOPO plasmid,
resulting in predominantly short (~50-100 bp) fragments of DNA with
ClaI-compatible overhangs (Fig. 2, lane
1). By ligating the ClaI-digested pseudogenome with an
excess of stuffer-adaptor DNA (1 : 30 molar ratio) we obtained high
molecular weight products of around 50 kb (Fig. 2,
lane 4) which were resistant to ClaI digestion,
indicating that the stuffer-adaptor DNA efficiently blocked
unmethylated ClaI sites from re-digestion (Fig. 2, lane 5). Indeed, even when we loaded 10-fold
more stuffer-adaptor ligated pseudogenome treated with ClaI we
saw no evidence of ClaI digestion (data not shown). Although our
results clearly suggest that there was no re-ligation of unmethylated
pseudogenome restriction fragments following ligation of
stuffer-adaptor at a 1 : 30 molar ratio, it would be feasible to
increase the ratio of stuffer-adaptor DNA to genome fragments, thus
decreasing the potential for unwanted re-ligation even further.

We suggest that the prolonged ligation we used resulted not only in the
concatamerization of restriction fragments but also in the
self-ligation of concatamers. Consequently, rolling circle
amplification by phi29 was possible, ensuring that long fragments of
DNA (>40 kb) were generated during subsequent amplification and thereby
attenuating the loss of restriction sites.

After the concatamerization reaction, the mixture was subject to phi29
amplification. The highly viscous nature of the phi29 solution
following amplification and the fact that most DNA remained immobilized
in the loading well of the 0.6% agarose gel we used supports the high
molecular weight of the DNA obtained (Fig. 2, lane
6). Subsequent digestion of the phi29-amplified DNA with
ClaI led to the excision of a 6-kb fragment corresponding to the
region between the originally methylated ClaI sites of the
plasmid (Fig. 2, lane 7). Thus, although the
original unmethylated fragments are carried through the procedure,
their ClaI sites are rendered indigestible, thereby negating
contamination of the methylated DNA pool.

Our method should therefore be of use in performing scans for
differential methylation across an entire genome. Moreover, coupled to
next-generation end-sequencing technology, a digital readout of the
prevalence of methylation marks at a particular locus can be
ascertained — of particular value to complex tissues such as
plant callus or to comparisons of different organs or developmental
stages.

Although we have used ClaI here, by modifying the sequence of the
stuffer-adaptor DNA, any methylation-sensitive endonuclease could be
used. Thus, the granularity of genome methylation scans can be adjusted
to suit the research agenda, with rare-cutting endonucleases producing
a coarser scan at reduced costs with increasing frequencies of cutting
producing finer maps at greater cost. Finally, our method circumvents
potential difficulties in aligning read sequences from
bisulfate-treated samples (which ultimately convert unmethylated C to
T) to a genome. This is particularly pertinent in plants where perhaps
close to half of methylation marks exist not only in a CpG context, but
also in CHG and CHH contexts (where H refers to A, C, or T) [5].

This work was supported by an Australian Research Council Centre of
Excellence grant (CEO348212) to the University of Newcastle Node of the
Centre of Excellence for Integrative Legume Research to RJR and an
Australian Research Council Australian Postdoctoral Fellowship
(DP0770679) to MBS.